While my previous blog covered the need to separate EtO from the air peaks introduced into the instrument, there are several other potential interferences that have to be managed. Acetaldehyde has an almost identical structure and mass spectrum. It is very common in nature and produced in a wide variety of industries, and it is possible to be present in both lab blanks and samples. Methanol also shares several ions with EtO, and as a common solvent for volatile standards (e.g., 8260 and TO-15A internal standards) it’s likely present in most air labs.

Fig. 1 – Comparison of EtO, Acetaldehyde, and methanol mass spectrum

Fortunately the cryo cooling helps with these separations as well. In addition, I also found that using selected ion monitoring (SIM) produced a cleaner baseline and better signal to noise ratio, allowing for detection of EtO down to 0.05ppb or lower (Fig.2 lower trace).

Once the troublesome EtO/acetaldehyde/methanol separations are solved with cryo cooling, I was able to use EZGC to get a working oven program to separate the TO-15A compound list. Without the need for extra sensitivity on the TO-15A compounds I found it helpful to use the combined SIM/Scan capabilities of the Agilent 5977A mass spec, using the SIM data for EtO and the scan data for the TO-15A list. This meant I didn’t have to optimize the SIM parameters for nearly 80 compounds, keeping the method much simpler.

While many labs may be reluctant to use cryogenic cooling due to costs and safety issues, it can be a powerful tool to separate out very volatile compounds. Here it was critical in the separation of EtO from methanol and acetaldehyde. In addition, the ability to acquire both SIM and scan MS data allowed for the increased signal to noise ratio for EtO in SIM mode, while maintaining the simplicity full scan for the TO-15A compounds. Together, cyro cooling and SIM/Scan can allow for the relatively simple addition of EtO down to 50ppt to TO-15A analysis.

“One last talking point: We told you there is no free lunch when incorporating the SPME Arrow into your laboratory, because you will have to install a GC Inlet Conversion Kit. It is important to stress that the inlet conversion kits work with all 3 types of SPME Arrow AND you may use the converted inlet with all your standard injection techniques (e.g., liquid syringe, headspace syringe, etc.). So, there is no need to swap back and forth between inlets.”

What I would like to do in this post is to help you select the correct Conversion Kit for your specific model GC. First, here again is a link to the Conversion Kits which we sell.

The table may look a little intimidating for those not familiar with it, so let’s break it down into more manageable sections, but before we do that, listed below are a few things to keep in mind.

A. Each kit is designed only for a split/splitless injection port for the particular GC make/model which is listed in the product’s description. If your GC has two split/splitless injection ports, you can install a conversion kit onto either or both split/splitless injection ports. However, these conversion kits are not for on-column, LVI, PTV or any other type of injection port.

B. Each kit is designed only for the specific GC make/modellisted. While it is possible that a kit may fit a different model of GC, since we do not have every make/model of GC at Restek, we can only guarantee that it will fit the model(s) listed.

C. Please read the description of the kit very carefully since there may be additional stipulations for a proper fit. As an example, for the Agilent 6890 Conversion Kits, the split vent filter needs to be the type which is referred to as the “canister-type filter”. Here is a photo of this canister-type filter.

D. Each instrument has three options for the conversion kits.

1. A kit without any additional products from Merlin Microseal. These kits should include all you need to get started with either the 1.1mm or 1.5mm Arrow fiber.

2. A kit with additional products from Merlin Microseal for 1.1mm Arrow fibers.

3. A kit with additional products from Merlin Microseal for 1.5mm Arrow fibers.

E. As you may have guessed, you should not attempt to use a 1.5mm Arrow fiber with a kit designed for a 1.1mm Arrow fiber. In addition, we do not recommend that you use a 1.1mm Arrow fiber with a kit designed for a 1.5mm fiber because you will not get proper sealing around the SPME Arrow. To further clarify, each Merlin Microseal has a specific “duck bill” designed to tightly seal around that specific SPME Arrow diameter. You may wish to review the following post for additional information on how the Merlin Microseal works.

F. As Jason mentioned, once any conversion kit is installed, you do not need to uninstall the kit to use regular SPME fibers (you can also keep using your same syringe). However, you may want to replace the injection port liner with a more appropriate internal diameter liner. For example, if switching from an Arrow fiber to a regular fiber, a smaller ID liner is usually used, such as 0.75mmID (or 1.0mmID).

Below are the various GC makes/models in which Arrow Conversion Kits are available. I hope this post has helped simplify the selection of Arrow Conversion Kits which Restek offers. Please let us know if you have any questions.

Split and splitless are the two most common modes of injection for GC. Each has its own set of pros and cons, with the required method sensitivity vs the sample concentration ultimately dictating which one to use. For trace level analyses, a splitless injection is often the best choice, as the goal is to recover close to 100% of everything that is injected into the instrument. Unfortunately, splitless injections can be tricky to optimize. There are a number of parameters that must be carefully adjusted to ensure the best chromatography possible.

During a splitless injection, the split vent is closed, directing all flow to the column (with the exception of the septum purge). Because of this, total inlet flows are relatively slow, leading to slower transfer of the analytes to the column compared to split injection. This can lead to diffusional band broadening (wider peaks), as well as increased time for adverse interactions to occur within the inlet.

The first critical decision one must make when performing splitless injections is selection of an appropriate inlet liner. I have discussed liner selection in detail in a previous blog series, so I will refer you to the following blogs for detailed information pertaining to splitless liner selection:

Beyond the liner there are several instrument settings that must be optimized to ensure successful splitless injections. These include inlet temperature, splitless hold time, and initial oven temperature. In the upcoming blog series, I will discuss the importance of each of these parameters, what can go wrong if they are not properly optimized, and some tips for optimizing them for your analysis.

To continue my blog parts 1 & 2 (Part 1:https://blog.restek.com/ p=67087,https://blog.restek.com/chiral-separation-on-a-c18-column-separation-of-d-and-l-amphetamines-part-ii/) , where I have discussed the importance of separating the chiral d- and -l isomers to accurately identify the illicit isomer using an achiral method on a Raptor C18 column employing a pre-column derivatization technique. Today I’d like to discuss more about the sample derivatization procedure and optimization of the derivatization conditions, which is the main step for separation of the chiral compounds on a reverse phase C18 column.

When it comes to sample derivatization, we think about how can we get the best derivatization conditions for our workflow. As a chemist, we want a procedure that’s simple, fast and reproducible with maximum derivatization efficiency and easy to replicate in other labs.

In the current published methods, samples like urine and oral fluid employ 2 steps sample preparation: first the biological samples are cleaned up using solid phase extraction, followed by derivatizing the sample to separate the chiral amphetamines and methamphetamines to detect the presence of illicit isomer.

Novelty of this application: So, how is this work different from the published or existing procedures for the separation of chiral compounds on a reverse phase column? Here, we employ one step sample prep procedure: that is direct derivatization of urine sample without any cleanup, followed by evaporation and reconstitution with the starting mobile phase of the LC-MS/MS gradient conditions, where the reconstitution serves as dilution of the urine sample, to minimize the matrix effects. Sample centrifugation followed by filtration using filter vials also helps in mitigating the matrix effects.

Marfey’s Reagent: FDAA is 1-fluoro-2-4-dinitrophenyl-5-L-alanine amide, also called Marfey’s reagent. This reagent reacts with primary amines of the enantiomers or chiral compounds and convert them into diastereomers and are detected using UV and MS/MS in liquid chromatography methods. FDAA derivatives of D-isomers exhibit strong intramolecular bonding, which reduces their polarity relative to the corresponding L-isomer derivatives. Consequently, the D-derivatives are selectively retained on reverse phase columns and elute much later than corresponding L-derivatives.

Sample Derivatization Procedure: 50 μL of calibration standard or QC urine sample was aliquoted into a microcentrifuge tube. 10 μL of a working internal standard (20 μg/mL (±)-amphetamine-D11 and (±)-methamphetamine-D11 in water) and 20 μL of 1M NaHCO₃ was added and vortexed at 3000 rpm for 10 seconds. After vortexing, 100 μL of 0.1% (w/v) Marfey’s reagent (1-fluoro-2-4-dinitrophenyl-5-L-alanine amide) prepared in acetone was added, vortexed, and heated at 45 °C for 1 hour. Samples were allowed to cool to room temperature before the addition of 40 μL of 1M HCL in water. The sample was then vortexed and evaporated to dryness under nitrogen at 45 °C. Samples were reconstituted in 1 mL of 40:60 water: methanol (v/v) and filtered using Thomson SINGLE StEP standard filter vials (cat.# 25893) and then injected.

Optimization of Derivatization Procedure: In order to obtain the best sensitivity and to achieve maximum derivatization, a series of experiments were performed using a 4000 ng/mL d- and l-amphetamines and d- and l-methamphetamines sample prepared in water. To determine the optimal derivatization conditions, various incubation times and derivatizing reagent volumes were assessed. For the length of the incubation time experiment, samples were incubated at 45 °C with 100 µL 0.1% (w/v) Marfey’s reagent for 15, 30, 45, 60, or 90 minutes. For the volume experiment, varying volumes of 0.1% w/v Marfey’s reagent (25, 50, 100, or 200 µL) were added to the samples, followed by incubation at 45 °C for 60 minutes. All the samples were subjected to the sample preparation procedure previously described and all conditions were evaluated in quadruplicate.

As shown in Figure 1, the highest peak responses were obtained using 100 µL of 0.1% (w/v) Marfey’s reagent with a 60-minutes incubation time at 45 °C. Longer incubation times showed decreased peak area responses for both amphetamines and methamphetamines diastereomers. Therefore, 60-minutes incubation time was selected for this method.

Figure 1

Figure 2. depicts that increased concentrations of the derivatizing reagent negatively affects the analytes sensitivity, 100 µL of 0.1% (w/v) Marfey’s reagent incubated at 45°C for 60 minutes resulted in the highest signal response compared to both higher and lower levels of the Marfeys reagent for all the analytes.

Figure 2.

Incubation: Water Bath or a hot plate? I would recommend using a water bath, as the temperature distribution was found to be even to the sample tubes from the water compared to an oven or a hot plate.

Concentration of derivatizing reagent: Do not to use the derivatizing reagent in excess concentrations, excess derivatizing reagent in the samples can negatively affect the analyte signal sensitivity and mass spectrometer ion source.

Always prepare the reagent fresh in Acetone and store in a dark place. Based on the above discussed results, these optimized derivatization conditions for both derivatizing reagent volumes and incubation times were used for all subsequent experiments and method validation in urine.

Method development for Quechers sample cleanup can be a complicated task. Not only does the analyst need to make sure their analytes of interest can be recovered, but the matrix interferences must be removed enough to make the quantitation possible and reliable. It makes sense that selecting the best dSPE products is highly dependent on the specific matrix that is in the sample.

If you are not familiar with Quechers techniques in general, I highly recommend a couple of videos that we have here in our Restek library to get started:

After you complete the 1st part of the Quechers procedure, which is extraction with a Quechers salt, you will need to take a subsample (aliquot) of the supernatant from that extraction to apply sample cleanup with dSPE. If you are lucky, there may be a validated method an analyst has performed and published that you can use for as a guide for your analysis. Even so, you will need to make sure the conditions are optimized. Usually if there is no existing method to follow, the analyst needs to develop a method for their specific combination of analytes and sample matrix.

SIZE

The first issue to consider is which size of Quechers tube you need for dSPE, since we sell the sorbents in smaller quantities in 2 mL centrifuge tubes or larger quantities in 15 mL tubes. These are designed for extract aliquot volumes of 1, 6, or 8 mL. If you are only analyzing the final extract one time with one analysis method, a small aliquot volume of 1 would be fine to use. This would call for using the 2 mL tube. If you will need to split the final extract for 2 or more analyses, a sorbent mixture in a 15 mL tube, designed to process aliquot volumes of 6 or 8 mL, will give you more extract to work with. The following table from our instruction sheet shows the aliquot volumes for which each of our dSPE products are intended.

WHICH SORBENT

Selecting the best sorbents to use depends on what you need to remove from your sample. Removing interferences is only about two thirds of the battle, though. You also need to determine which combination of sorbents will remove the interferences from the matrix AND still leave your analytes intact enough to give good recoveries. Here is a link to our listing of dSPE products on our website: https://www.restek.com/catalog/view/7186

To optimize the removal of interferences, you can select sorbents as follows to fit your sample matrix characteristics.

To remove fats/lipids and waxes- use C18 (endcapped C18 bonded to silica)

To remove pigments- use GCB (graphitized carbon black)

The above listed are the main categories, but be aware there is some crossover, because each sorbent has secondary interactions also that may come into play. (Be on the lookout for more discussion of these in the next blog post.)

HOW MUCH SORBENT

Aim high or low for each sorbent, depending on the levels of each type of interference.

For example, if your sample is a vegetable, it may contain a low amount of sugars or acids. Therefore, I would start with a minimal amount of PSA.

Another example-if your sample does not contain much pigment- let’s say it is something like celery. In that case, you probably do not need GCB, but you may need a small amount of C18 because there will be a bit of waxy material and cellulose. PSA will help with the cellulose also.

A 3rd example- your sample is highly pigmented and a high amount of sugar, like red grapes. You will need PSA and GCB in your sorbent selection. You may need a small amount of C18 as well, but you could try without it to see which way works better.

HOW TO MAXIMIZE RECOVERY OF ANALYTES

The good news is that most of the dSPE sorbents are safe for pesticides because that is what Quechers was originally developed for. There is one exception- GCB does have the potential to interact with planar nonpolar molecules. There are a few pesticides that are planar and prone to such interaction. If those are important targets in your analysis, you might consider not using GCB or keeping it to a minimum. If you are analyzing something like PAHs (Polynuclear aromatic hydrocarbons) instead of pesticides, keep in mind that most of those compounds are planar and would have a strong interaction with GCB. Most of the successful Quechers methods used for PAHs use a minimal amount or no GCB.

The final determination for sorbent selection should be made by experimentation and method development. Once you arrive at a combination or sorbents that you think might work best, you can perform a method validation to ensure reliability and ruggedness. (Validations include processing a set of control samples that are fortified with analytes prior to extraction to mimic levels that one might encounter for real samples. The concentrations and number of replicates required are usually determined by regulatory guidelines or protocols.)

The growing variety of cannabis products that are currently available in the market speaks volumes about the creativity of entrepreneurs within the industry. Vapes, lotions, balms, cookies, hard candy, gummies, infused drinks, chocolate, fudge, soaps, and tinctures are only a handful of the product types that are currently being commercialized in this ever-growing market. Such explosion of marketing creativity in launching innovative cannabis goods to satisfy the needs of different consumers, demands resourcefulness from analytical chemists in finding smart and reliable solutions to comply with regulations in different countries/states. For instance, in the majority of states where cannabis has been legalized, potency testing (analysis of cannabinoids) is required in plant material, concentrates, and in all sorts of final products to ensure that the label claims truly match the product composition. Given the high concentration of cannabinoids in any cannabis commodity, HPLC-UV is the instrumental technique of choice; however, special attention should be paid to the sample preparation process to ensure close to exhaustive recoveries from any matrix. Testing pesticides and mycotoxins is a completely different story. Analysis of these compounds is normally required at part per billion (ppb) levels, which demands the use of more sophisticated equipment, namely liquid chromatography-tandem mass spectrometry (LC-MS/MS) and gas chromatography – tandem mass spectrometry (GC-MS/MS). In the majority of the states and in Canada, analysis of pesticides and mycotoxins is obligatory only in plant material. However, the state of California, which is known for having some of the most rigorous cannabis regulations, demands testing for these contaminants in any cannabis good. To support California cannabis testing, we recently published a technical article on the analysis of pesticides and mycotoxins in brownies that you can check here. In that article, we described the different steps we followed to optimize the extraction of the California list of pesticides and mycotoxins using brownies as a model matrix. Based on this work, on my recent experience working with dark chocolate, and after talking to several analysts from Cannabis testing labs that still have concerns about how to properly quantitate pesticides and mycotoxins, I decided to write this series of blogs to highlight some important points, critical for performing quantitation of these contaminants in any matrix. The first point I would like to discuss is matrix effects.

Matrix effects

To extract all the target pesticides and mycotoxins from any cannabis matrix, solvents like acetonitrile are commonly used. However, such extraction conditions will also favor the extraction of multiple matrix components, which will vary depending on the sample type. Co-extracted matrix components can have an effect on the MS/MS response of target analytes. That effect is what is collectively known as matrix effects. In the case of LC-MS, both electrospray (ESI) and atmospheric pressure chemical ionization (APCI) are susceptible to matrix effects (ESI is more prone to matrix effects than APCI). So, how do you know if your methodology is being impacted by matrix effects? One of the best ways to evaluate matrix effects is by following the approach proposed by Matuszewski et al. [1,2]. In this approach, you compare the responses of your analytes post-spiked in blank matrix extracts versus their response in neat solvent by using the following equation:

In an ideal world, your result should be 100. Values below 100 indicate that you have signal suppression, and values above 100 show signal enhancement. To illustrate the importance of performing this test in each type of matrix that you have to analyze, we can refer to Figure 1.

Figure 1. Matrix effects corresponding to daminozide in brownies, dark chocolate and cannabis biomass extracts. Brownies samples were prepared as described in our technical article (here). Dark chocolate samples (0.5 g) were extracted by using isopropyl alcohol (0.5 mL) and acetonitrile acidified with acetic acid at 1% (2.5 mL); subsequently, 2 mL of the extract were passed through a Restek Resprep C18 cartridge (Restek Cat.#26030). Cannabis biomass was prepared by adding 6 mL of acidified acetonitrile to 1g of sample; the final extract was passed by a Restek Resprep C18 cartridge (Restek Cat.#26031) (cannabis biomass experiments were conducted at Convergence Lab facilities in Santa Rosa, CA). In all cases, extracts were mixed with water in a ratio 3:1 (extract:water), and 2 µL were injected in the LC-MS/MS system.

As can be seen, the same pesticide, daminozide, analyzed under the same chromatographic conditions displays different levels of ionization enhancement in brownies (209%) and chocolate (175%) while, a certain level of ionization suppression is observed in cannabis biomass extracts (76%). Since this compound is highly polar and therefore poorly retained under reversed phase conditions (retention time = 0.7 min), it co-elutes with multiple co-extracted polar interferences that can affect its ionization in matrix extracts. Unfortunately, improving chromatographic retention in the same method for the analysis of all the California regulated pesticides and mycotoxins is difficult as most of the compounds are well analyzed via reversed phase chromatography. Typically, when you are performing multiresidue pesticides testing, compromises need to be made for certain analytes. Now, you may wonder what are the best conditions to assess matrix effects once you have a potential analytical workflow ready? Here are some tips.

First, check your matrix blanks. The surrogate matrix that you choose to develop your method may contain some of your target analytes. Ensuring that your blanks are true blanks is important not only for the assessment of matrix effects, but also for method development in general. If a matrix blank is not easily available, in some cases, subtracting the area of the blank from the post-spiked blank extract could help to assess matrix effects.

Check your MS/MS transitions in different samples (matrix blanks vs. post-spiked extracts vs. standards). Although this step is not directly related to evaluating ionization effects, it is always important to make sure that you don’t have interferences sharing the same transitions of your target analytes and eluting at the same retention time. If this is the case, picking another transition with higher selectivity for the detection of your target analyte, even if it isn’t the one with the highest intensity, is the best choice (you may want to check the blog written by my colleague Dan Li here).

Evaluate matrix effects at different concentration levels. If you pick a concentration that is too high to investigate matrix effects, your results may indicate that your method is free of ionization suppression/enhancement. However, evaluation at levels close to the LOQ or even at the requested action level, ionization effects may become apparent. In our brownies method, we spiked extracts at 15 ng/mL to assess matrix effects. Why? If we take into account that we used was 0.5 g of sample, 3 mL of solvent for the extraction, and that the lowest action level in the California list of pesticides is 100 ng/g, in the case of having close to exhaustive recoveries we were expecting analytes to be at a final concentration of 16.7 ng/mL in sample extracts. Based on this, we considered it appropriate to choose a concentration close to 16.7 ng/mL. Nonetheless, it is important to highlight that the evaluation of matrix effects at levels close to the LOQ is always recommended.

Use the average response of at least three replicates. Technical replicates are critical to ensure that you have a representative value with an estimated error.

Make sure that your neat solvent composition matches your final extract composition. In our case, we used acidified acetonitrile to prepare the neat solvent samples to assess matrix effects in brownies extracts (you can check our technical article to see why we used acified acetonitrile).

So, what if I have matrix effects? Matrix effects are very common, and that is why we recommend the use of a matrix matched calibration to account for variations in the ionization conditions during the entire chromatographic run. In cannabis testing it can be difficult to perfectly match the sample that is being analyzed due to the broad variety of products. However, we recommend trying to find a surrogate matrix very close to what you are analyzing. It is worth emphasizing that the use of neat solvent calibration solutions can lead to biased results. For instance, in the case of daminozide, quantitation in brownies and chocolate (Figure 1) using calibration solutions in solvent will cause an over-estimation of the analyte concentration. In addition to matrix-matched calibration, the use of isotopically labeled analogues as internal standards is essential. Although we understand that this represents an extra cost that many people would prefer to avoid, internal standards are necessary to quantitate your target analytes in the great majority of cases, and especially when dealing with complex matrices. Selecting a handful of representative deuterated analogues that elute at different retention times in the chromatographic run is the best way to go when dealing with multiple analytes. In the case of daminozide, we had to introduce its deuterated analogue to account for ionization effects at the right retention time as well as to account for low absolute recoveries (this story will be part of our next blog). If your matrix effects are significantly affecting the detection of your analyte of interest at the required levels, you may need to play with your chromatographic method to separate your target analytes from interferences, or you many need to adjust your sample clean-up conditions.

What about GC-MS/MS analysis? Are there any matrix effects?

Indeed, co-extracted interferences can also affect analytes’ responses in GC-MS/MS analysis. Co-injected and non-volatile matrix components can interact with active sites in the GC column or injection port, and this causes an alteration in the response of analytes injected in matrix extracts in comparison to neat solvent standards. This effect is known as “matrix-induced chromatographic response enhancement” and it is particularly problematic for compounds that are polar and/or thermally sensitive [3,4]. The non-volatile matrix components that are co-injected help to block active sites in the GC system, hence preventing adsorption and thermal degradation of analytes. This leads to an enhancement of the analytes’ response in comparison to neat solvent injections where broader peaks with lower responses can be observed. Although this may sound as if having co-extracted matrix components could be beneficial, having appropriate sample clean-up is crucial to avoid matrix built-up in the GC system after multiple sample injections [3]. Running extracts that are too dirty can lead to losses in analyte response, peak tailing, and eventually MS system contamination.

The best way to prevent “matrix-induced chromatographic response” effects from leading you to biased quantitative data is by using matrix-matched calibration. For this reason, in our brownies method we used the same extract for both LC and GC analysis (for GC injections we did an extra clean-up step). Alternatively, the use of certain compounds capable of blocking GC active sites, also known as analyte protectants, has been proposed. Although we haven’t tested this strategy yet for the analysis of cannabis products, it is definitely in our to-do list!

I hope this blog was helpful for your cannabis method development work! Please stay tuned for part II. If you want to do some extra reading on this topic, here are some of the cited references:

Per- and polyfluoroalkyl substances (PFAS) are ubiquitous in our society and can be found in a wide range of consumer products; hence they are widely present in the environment, people and animals. They are persistent chemicals that have the potential to accumulate. Therefore, they are evaluated and monitored using a variety of methods and controlled by numerous regulations. Due to its every-where presence, it is no wonder they end up in our samples, systems or chemicals elevating our results and making us waste numerous hours searching for contamination to get the analysis to pass. Where do they come from? What are some of the most overlooked sources of contamination? How to avoid those false positives?

To abate the cross-contamination with PFAS, follow your sample journey from the sampling to the chromatogram on your screen. Just assure the samples never come in contact with material made or, coated with PFAS, which can be often easier said than done.

Instead, for sampling use HDPE (high-density polyethylene) and polypropylene bottles and label the sample after the sampling. Some labs offer pre-screened sampling bottles.

Sample preparation:

Avoid glass transfer pipettes, vial caps with PTFE seals and even some glass HPLC vials (PFAS has been known to absorb to glass when it has been in contact for an extended period of time. Glass bottles or containers can be used if they are known to be PFAS-free). Who would have thought aluminum foil has a nonstick coat? Avoid wearing sunscreens/insect repellents, fabric softener, moisturizers and lab coats (stain-resistant). Avoid washing glassware with detergents that haven’t been tested for PFAS presence.

An effective tip for all methods using SPE manifold is after analyzing SPE samples, all reusable parts should be washed with acetonitrile and sonicated for ~10 minutes to avoid cross-contamination.

Instrumentation:

Avoid all PTFE tubing, PTFE frits, and sample valves with certain rotor seal or stator face. Be aware that when your LC-MS/MS system has been sitting overnight or extended time frame the first few injections will show a high level of PFAS due to the buildup of residual PFAS in the system.

Instead, replace all tubing with PEEK tubing or stainless steel in your system to avoid system-related interferences. You can find kits specifically designed for PFAS analysis from individual instrument companies (sometimes this can be costly and not feasible).
A great option is to add a PFAS Delay Column (Cat.#: 27857) after the mixing chamber to trap system-related PFAS. See references listed at the bottom of this blog for PFAS Delay column video and another blog explaining it in greater detail. Before starting the analysis condition your analytical column (Raptor C18, (Cat.#: 9304A52) for methods such as US EPA 533, US EPA 537.1, ISO 21675:2019, ASTM D7968-17A, ASTM D7979-19, DIN 38407-42 and many more) with 20-30 column volumes. Refer to our PFAS column selection guide listed in the reference for additional dimensions. Execute 3-4 injections of high organic solvent (acetonitrile) while running your method to flush out any residual PFAS in your injector.

Solvents:

A part of the system that is often overlooked when searching for PFAS contamination is the mobile phase. Many HPLC and LC-MS/MS grade mobile phases (water, acetonitrile, and methanol) use PTFE filters before they are bottled by the company. Bottle caps could have PTFE lining.

If contamination is found in the brand new solvent bottles, filter all the HPLC or LC-MS/MS grade water with a PFAS Delay column (Cat.#: 27857) before you make the buffer. Lot check all solvents and stick with a lot that works.

The most influential part to avoid contamination in PFAS analysis is to have a designated system, mobile phase, and materials that you know are PFAS-free. Any time you introduce a new part/solvent to the analysis test to make sure it is clean of PFAS. Good lab practice is always your friend.

In Part I of this series, I posed the question of the impact of the amount of wool on liner performance. As a reminder, this study specifically examines Topaz liners that are prepacked with wool and then deactivated. With your own hand-packed liners these results will not apply.

The first criteria I wanted to examine was inertness, as it’s generally the largest concern when using wool. Since wool has a high surface area, it can be difficult to thoroughly deactivate, leading to the theory that the more wool, the more activity. While this is likely true if you’re packing your own liners, the results for the Topaz liners in this study, where the wool was deactivated in place, did not show this trend.

Average response factors for active acids and bases injected splitless at 0.5 ng on different liners can be found in Figure 1:

Figure 1: Average relative response factors for several active acids and bases for each group of liners, organized by weight-length of wool plug. Non-reactive compounds are included for comparison. Error bars represent 1 standard deviation.

As you can see from the above results, differences in relative responses for most of these active compounds were small, even with liners that were overpacked (10 mg and 15 mg of wool). On the other hand, for 2,4-dinitrophenol, an often-troublesome acidic compound, the liners that were packed with 15 mg of wool showed a significant decrease in response. Interestingly, for two of the difficult to analyze bases, dicyclohexylamine and benzidine, response factors slightly increased as wool amount increased. Perhaps the wool has a slightly basic character.

The above data is presented using a response factor, n-tridecane as an internal standard. This allows us to examine differences caused by activity by normalizing to the internal standard. But what if we remove this normalized data and see the general effect on vaporization? Figure 2, below, illustrates raw peak area for the acids and bases for the different liner groups.

Figure 2: Average peak area counts for several active acids and bases for each group of liners, organized by weight-length of wool plug. Non-reactive compounds are included for comparison. Error bars represent 1 standard deviation.

Notice that the overall trend is that peak areas tend to increase as wool amount increases, though the differences are generally pretty small. This could be due to the enhanced vaporization from the higher surface area of the wool. 2,4-dinitrophenol is an exception, as you can see the large loss in area when overpacked with 15 mg of wool.

Besides response factors and raw area response, we can also examine tailing factors of active compounds. Figure 3 shows a comparison of tailing factors for select active compounds. There were not any significant differences, even with the liners that are very overpacked with wool. I found this interesting for 2,4-DNP, since I did observe a reduction in response on the 15 mg packed liners, yet the tailing did not increase. This tells me that 2,4-DNP is likely experiencing irreversible adsorption within the wool, rather than reversible, which might be exhibited by increased tailing.

Figure 3: Tailing factors for select active compounds for each group of liners, organized by weight-length of wool plug.

In addition to inertness, I wanted to examine retention times of volatile compounds, which could be affected by flow differences or increased interactions with the wool.

The most volatile compound in this study was cis-1,2-dichloroethene. As everything else in the system was left unchanged, any differences in retention should be due to the differences in wool packing. The average observed retention times for each group of liners is shown in Figure 4 below.

Figure 4: Retention times of cis-1,2-dichloroethene for each group of liners, organized by weight-length of wool plug.

The liners with more wool had more retention of cis-1,2-dichloroethene. For the liners packed with 15 mg of wool, this volatile compound eluted around 4 seconds later than the liners that were packed within specifications. While differences are seen when packed far beyond the specifications, liners at the upper and lower specs for wool did not show significant variation in retention time.

If we look at the most volatile compound from the active acids and bases mix, 4-picoline, we can see a similar trend (Figure 5). The liners with 15 mg of wool showed a 6 second increase in retention for 4-picoline.

Figure 5: Retention times of 4-picoline for each group of liners, organized by weight-length of wool plug.

One final observation was with regards to liner “bleed”. Generally, upon installing a new liner, you may notice some peaks that are related to excess deactivant “bleeding” off of the liner, especially during the first injection or two. In general, I have observed very little bleed coming off of Topaz liners. I did observe, however, that the liners that were packed at 2 or 3 times the weight specification for wool showed increased bleed on an initial solvent blank (Figure 6).

The above data demonstrates that for Topaz liners packed within the specifications, there are no major differences in performance based on the amount or density of the wool. When packing at 2-3 times the maximum weight specification, there were some differences observed: 2,4-DNP response was negatively affected, retention time shifts occurred, and there was increased liner bleed. I believe the lack of differences observed in many cases is owed to the deactivation process itself, with the wool being deactivated in place using vapor deposition.

I don’t want to sound like a broken record, but I’ll again point out that results may look completely different if you are packing your own liners. This is due to breaking fibers and exposing active sites during the process. If anything, this data should help to further convince you to try prepacked liners with the wool deactivated in place, since in addition to superior inertness, you will get better reproducibility.

Sometimes we get asked by customers for clarification when installing/connecting a new or replacement weldment from their purge & trap autosampler to their Agilent GC’s split/splitless injection port. As a result, I wrote this post to assist those that may be new to this type of installation or those that simply could use a quick refresher on the topic.

Here are the related products we sell (links below) which will accomplish this task. I had chosen item # 22664 as the product for this example, but the instructions (shown below) should also suffice for the different purge & trap replacement weldments we offer.

2. Connect the two gas lines show by the green arrow in the photo below. Make sure the lines are installed properly. The line labeled as “C” represents carrier gas and the line listed as “P” represents purge gas.

3. Unscrew the nut shown by the red arrow. Connect to the appropriate gas supply fitting on the purge & trap unit. Although a “jumper line” may be needed, this is unlikely (see bottom photo).

4. Connect the transfer line from the purge & trap unit onto the union as shown by the blue arrow. The transfer line is the carrier gas line exiting from the center of the heated jacket.

The other day, I and some of my colleagues started a discussion about how chromatographers are handling the current Covid-19 situation. Some are working from home, and some are experiencing temporary closures. Those that are working are likely working under “new” conditions – social distancing, additional personal protective equipment, possibly working different shifts, etc.

Since the tech support team is in our home offices, we use Skype and email to correspond. That has been a big change to us and likely a change for many of our ChromaBLOGraphy readers. As we talked, my colleagues immediately turned towards shutting down laboratory equipment. Laboratory equipment is expensive and it should be properly shut down to help ensure that it can be back up and running when needed.

• Go through your reagents and reference standards. Thoroughly check expiration dates. That is usually the first place an auditor focuses when visiting a lab. You can also make sure your inventory is accurate.

• Perform instrument qualification, if you are required to perform IQ/OQ/PQ in your industry. That way your instrument does not need to go off line, when you are in the middle of a project.

• Turn to Restek and read ChromaBLOGraphy posts. There’s a wealth of information.

• Maybe it has been a while since you had your ProFLOW 6000 calibrated, so have that completed. Restek recommends a yearly calibration to maintain its accuracy. Although the Leak Detector does not require a yearly calibration, it can be serviced/repaired to help maintain its functionality.

• Although not chromatography related, you can always start a home project, get out for a walk, or turn to a hobby. Personally, I’m going to need a new fly box (or two) to hold all the flies I have tied for fishing.